Chromobacterium subtsugae 
In 2000, a purple-pigmented bacterium (PRAA4-1) was isolated from forest soil in Maryland (Martin et al., 2004). In initial screens, this bacterium was found to be toxic to Colorado potato beetle and other insect pests (Martin et al., 2007a). Additional work with the isolate revealed activity gainst mites, grubs, diverse beetle species, aphids and plant parasitic nematodes, among other plant pests (Martin et al., 2007b, US Patent Application Publication No. 2012/0100236 A1).
Proteases and Insect Control
Proteases have the ability to target and destroy essential proteins and tissues of insects. Plants have naturally evolved to express proteases to protect against insects. Insect predators also produce protease in their venom, which contributes to mortality. Proteases have been identified as important insecticidal agents for control of insects in agriculture.
Proteases with insecticidal activity fall into three general categories: cysteine proteases, metalloproteases and serine proteases. Proteases of these classes target the midgut, cuticle and hemocoel. The peritrophic matrix of the midgut is an ideal target for insect control because it lines and protects the midgut epithelium from food particles, digestive enzymes and pathogens; in addition to acting as a biochemical barrier (Hegedus at al., 2009). Enhancins are zinc metalloproteases expressed by baculoviruses that facilitate nucleopolyhedrovirus infections in lepidopterans (Lepore et al., 1996). These proteases promote the infection of lepidopteran larvae by digesting the invertebrate intestinal mucin protein of the peritrophic matrix, which in turn promotes infection of the midgut epithelium (Wang and Granados, 1997). Homologs of enhancin genes found in baculovirus have been identified in the genomes of Yersinia pestis, Bacillus anthracis, Bacillus thuringiensis and Bacillus cereus (Galloway et al., 2005; Hajaij-Ellouze et al., 2006).
Plant cysteine proteases also demonstrate activity against lepidopteran larvae. Cysteine proteases in the latex of the papaya and wild fig trees are essential in the defense against various lepidopteran larvae. Toxicity to the larvae was lost when the latex was washed or when the leaves were treated with a cysteine protease-inhibitor, indicating that the defense may be due to the high concentration of cysteine proteases in the latex (Konno et al., 2004).
Proteases that target the cuticle are also important in insect control. The cuticle covers the entire outside of the insect as well as some invaginations of internal structures. The cuticle is composed of a waxy epicuticle, an exocuticle and an endocuticle that consist of protein, lipid and chitin (Harrison and Bonning 2010). Fungal infection of insects by Metarhizium anisopliae and Beauveria bassiana occurs when the fungal spores germinate on the cuticle, forming structures for penetration of the cuticle by a variety of enzymes, including proteases (Freimoser at al., 2003; Cho et al., 2006). One notable serine protease produced by M anisopliae, PR1A, digests the cuticle and plays an essential role in penetration (St. Leger et al. 1987). A clone of M anisopliae was engineered to contain additional copies of the py1a gene and showed 25% more kill of tobacco hornworm than the wild-type (St Leger et al., 1996). B. basianna was also engineered to express the M. anisopliae PR1A protease and demonstrated increased toxicity of larvae of the Masson's pine caterpillar, Dendrolimus punctatus, and the wax moth, Galleria mellonella (Lu et al., 2008).
The basement membrane of insects consists of proteins that surround the tissue and contribute to a variety of functions from structural support to barriers for viruses. Three potential basement membrane-degrading proteins were evaluated using Autographa californica multiple nucleopolyhedrovirus (AcMNPV). This baculovirus was engineered to express two vertebrate metalloproteases, rat stromelysin and human geatinase A, as well as the fruit fly cathepsin L, ScathL. The ScathL protease demonstrated the best baculovirus activity. The median survival time of infected tobacco budworm larvae was reduced by 50% when compared to wild-type infected larvae (Harrison and Bonning, 2001). This data supports the idea that proteases expressed in viruses have the ability to access the basement membrane of insects, which generally functions as a barrier to viruses. A previous report identified two basement membrane proteins of imaginal discs of fruit fly larvae that are susceptible to hydrolysis by cathepsin L (Homma and Natori, 1996). Purified ScathL protease was also toxic to a variety of insect pests when it was injected into the hemocoel. The purified protease demonstrated similar melanization, mortality and hemolymph protease activity in lepidopteran larvae as was seen ScathL expressed baculovirus infections (Li et al., 2008). Basement membrane damage is cause by purified ScathL protease both in vivo and in vitro (Tang et al., 2007; Philip et al. 2007).
Arthropod predators have also been shown to contain basement membrane cleaving proteases in their venom. One example is the parasitic wasp, Eulophus pennicornis, in which 3 metalloproteinases (EpMP1-3) were identified in the venom glands. Recombinant EpMP3 was injected into the hemocoel of Lacanobia oleracea larvae and resulted in significant mortality, or impaired development and growth in surviving larvae (Price et al., 2009). Social aphid soldier nymphs produce a toxic cathepsin B protease (cysteine protease) in their intestines. The protease is orally excreted into enemies and demonstrates insecticidal activity (Kutsukake et al., 2008).
A protease isolated from the bacterium, Xenorhabdus nematophilia, has been shown to suppress antibacterial peptides involved in insect immune response, making the insect susceptible to the pathogenetic process (Caldas et al., 2002). The enterobacterium, Photorhabdus luminscense, has been shown to be pathogenic to a broad spectrum of insects. The genome sequence of this bacterium identified genes related to toxicity, including proteases (Duchaud et al., 2003).
The use of proteases as insecticides has been of interest to plant modifications as well. Basement-membrane degrading proteases have been characterized and engineered for transgenic insecticidal protocols, with the goal of developing transgenic plants that are resistant to insect pests (U.S. Pat. No. 6,673,340, Harrison and Bonning, 2004). Proteases in the gut of insects have been shown to affect the impact of Bacillus thuringiensis Cry insecticidal proteins. Some proteases activate Cry proteins by processing them from a protoxin to a toxic form. Insect toxins have been modified to comprise proteolytic activation sites with the goal of incorporating this modification into transformed plants, plant cells and seeds. Cleavage of these sites by the insect gut protease results in an active insect toxin within the gut of the pest (U.S. Pat. No. 7,473,821, Abad et al., 2009).
Insecticidal Activity of Chitinases
Chitinases expedite insecticidal activity by puncturing the insect midgut lining and degrading the insect cuticle. Degradation of these membranes exposes the insects to pathogens, to other insecticidal compounds, and/or to plant defenses.
Chitinases hydrolyze the structural polysaccharide chitin, a linear homopolymer of 2-acetamido-2-deoxy-D-glucopyranoside, linked by β-1→4-linkages, which is a component of the exoskeleton and gut lining of insects. Chitinases are classified as either family 18 or family 19 glycosyl hydrolases. Family 18 chitinases are widespread, found in bacteria, plants, and animals; while family 19 chitinases are mainly found in plants (Henrissat and Bairoch, 1993). In insects, Chitinases play a role in molting (Samuels and Reynolds, 1993, Merzendorfer and Zimoch, 2003).
Chitinases alone show some insecticidal activity. Chitinase from Serretia marcenscens was found to be toxic to seventh instar Galleria mellonella larvae (Lysenk, 1976).
Transgenic plants which express insect chitinases have been shown to have increased resistance to insect pestss. Tobacco plants were transformed with cDNA encoding a Manduca sexta chitinase. Leaves from these transgenic plants were infested with Heliothis virescens larvae. After 3 weeks it was found that chitinase positive leaves had less larval biomass and feeding damage than chitinase negative leaves. It is possible that the activity of the chitinases render insects more susceptible to plant defenses (Ding, et al., 1997).
Insect cuticles provide a physical barrier to protect the insect form pathogens or other environmental hazards, and are composed primarily of chitin (Kramer, et al., 1995). Entomopathogenic fungi Metarhizium anisopliae, Beauvaria bassiana, Beauvaria amorpha, Verticillium lecanii, and Aspergillus flavus all secrete chitinases to break down the cuticle and enter the insect host (St Leger, et al., 1986, 1992, Campos, et al. 2005). According to Kim, et al., chitinase-containing supernatants of Beauvaria bassina were toxic to Aphis gossypii adults. However, when these supernatants were treated with an excess of chitin to inhibit the activity of the fungal chitinases, this mortality was significantly reduced, suggesting that chitinase plays an integral role in breaking down the cuticle and facilitating infection (Kim, et al. 2010). Chitinases have also been isolated from the venom of the endoparasitic wasp Chelonus sp., where they possibly help the venom penetrate the defenses of chitin protected prey (Krishnan, et al., 1994).
The peritrophic membrane, which lines the insect midgut, is another primarily-chitin-composed barrier that protects insects from pathogens. Any enzyme that can puncture this membrane has potential as a bioinsecticide (Wang and Granados, 2001). Hubner, et al. demonstrated that malarial parasites excrete chitinases to penetrate the peritrophic membrane in mosquitoes (Hubner, et al., 1991), and Shahabuddin, et al. confirmed that inhibition of chitinase with allosamidin is sufficient to prevent the malarial parasite Plasmodium gallinaceum from crossing the peritrophic membrane of Anopheles freeborni. Also, the addition of exogenous chitinase from Streptomyces griseus during the development of the Anopheles freeborni midgut prevented the formation of the peritrophic membrane (Shahabuddin, et al., 1993). This demonstrates that chitinases can break down the peritrophic membrane. Regev, et al. used E. coli to express Serratia marcescens endochitinase ChiA and confirmed with electron microscopy that Spodoptera littoralis larvae exposed to the endochitinase exhibited perforations in the peritrophic membrane (Regev, et al., 1996).
Because of the ability of chitinase to perforate the peritrophic membrane, endochitinases have also been shown to increase the insecticidal activity of Bacillus thuringiensis (Bt). Choristoneura fumiferana larvae reared on Agies balsamea treated with a mixture of a diluted commercial formulation of Bt and chitinase were killed more quickly than larvae reared on foliage treated with just Bt alone (Smirnoff, 1973). A mixture of a low concentration of Bt and S. marcenscens chitinase also resulted in higher mortality of Spodoptera littoralis larvae than Bt alone (Sheh et al., 1983). It is believed that this synergistic effect is due to puncturing of the peritrophic lining of the insect gut by the chitinase, facilitating the penetration of Bt spores into the insect. (Smirnoff, 1973).
Yen-Tc, an ABC type protein that is both necessary and sufficient for the entomopathogenicity of Yersinia entomophaga in the insect Costelytra zealandica, contains two family 18 chitinases, making it the first insecticidal toxin complex identified to incorporate chitinases. It is hypothesized that the chitinases are responsible for breaking down peritrophic membrane and exposing the midgut epithelial cells to the toxin. However, the chitinases may only be active in regions of the midgut with a relatively neutral pH (Busby, 2012).
Chitinases are also integral to the activity of some insect viruses. Hatwin, et al. created mutants of the Autographa californica nucleopolyhedrovirus (AcMNPV) that lacked the gene for chitinase. Usually, this virus causes liquefaction of the host larvae, facilitating the spread of the virus. This liquefaction did not occur when Trichoplusia ni larvae were infected with the chitinase negative virus. It was also confirmed that the AcMNPV chitinase is active under the alkaline conditions of the insect midgut (Hatwin, et al. 1997). A recombinant version of the same Autographa californica nucleopolyhedrovirus that expressed a Haemaphysalis longicornis chitinase was found to have bioarcaricidal activity against Haemaphysalis longicornis nymphs (Assegna, et al. 2006).
Rhs-Like Genes Encode Insecticidal Toxins
The rhs (rearrangement hotspot) gene family was first identified in E. coli. These genes confer chromosomal rearrangements by homologous exchange (Lin et al., 1984). They are 2 to 12 kb in size and exhibit a long core with a short tip. The core sequences are GC rich and highly conserved, but the tip sequences are GC-poor and highly variable. They encode proteins that have a large core domain and a short C-terminal tip domain. The protein core domain is hydrophilic and contains YD-repeats (Jackson et al., 2009). The Rhs proteins are capable of interacting with bacterial cell surfaces and binding to specific ligands (Wang et al., 1998). While the function of the Rhs proteins remains unknown (Hill et al., 1994), the structure is important because the YD repeats and highly conserved sequences resemble rhs and rhs-like genes encoding insecticidal toxins produced by bacteria.
Photorhabdus luminescens is a mutualistic symbiont of the nematodes from the Heterorhabditae family. The nematode infects the insect and injects the bacterium into the hemocoel of the insect. The bacterium then secretes toxins that kill the insect (Frost et al., 1997). Bowen et al. (1998), purified a high molecular weight protein associated with oral and injectable insecticidal toxicity that targets insects. In another study, Bowen et al. (1998) used high performance liquid chromatography to separate this protein into four toxin complexes (tc) termed, Tca, Tcb, Tcc, and Tcd encoded by the tc loci (Bowen et al., 1998). Waterfield et al. (2001) analyzed recombinant expression of the tc genes in E. coli to understand oral toxicity of Tc proteins. They found that without tccC-like homologs, they could not recover oral toxicity in E. coli. These authors concluded that TccC is involved in activation of toxin secretion. Furthermore, an amino acid sequence analysis revealed TccC and TccC-like proteins have a highly conserved core and highly variable extension. This structure bears resemblance to rhs-like elements (Waterfield N R, Bowen D J, Fetherston J D, Perry R D, and ffrench-Constant, R H, 2001). This similarity suggests that TccC-like and Rhs proteins share an ancient role in toxin mobility and activation for the Enterobacteriaceae family (ffrench-Constant, R et al, 2003).
Another microbe, Serratia entomophila, has insecticidal activity that targets New Zealand grass grub, Costelytra zealandica, and causes amber disease (Grimont et al., 1988). The virulence of S. entomophila is linked to a large plasmid called amber disease-associated plasmid (pADAP) (Glare et al., 1993). Hurst et al. analyzed the mutagenesis and the nucleotide sequence of pADAP to understand how it confers pathogenicity to grass grub. They found that pADAP encodes three genes responsible for the symptoms of amber disease, sepA, sepB, and sepC. All three genes are required for pathogenicity because a mutation in these genes abolishes amber disease. They illustrated that proteins encoded by the sep genes are similar to the proteins encoded by the insecticidal toxin complexes of P. luminescens. For example, the first 680 amino acids of SepC and TccC show a strong similarity. Furthermore, this region resembles the rhs elements of E. coli. The sepC gene is smaller than Rhs elements, but it encodes a hydrophilic protein core with nine Rhs peptide variants. Based on the similarity between the sep and tc genes, Hurst et al. concludes that these products are part of a new group of insecticidal toxins (Hurst et al., 2000).
Harada et al. discovered that, Pantoea stewartii ssp. DC283 is an aggressive pathogen that infects aphids (Harada et al., 1996). The aphid ingests the bacterium and DC283 is able to aggregate in the gut and cause death of the aphid. Stavrinides et al. performed a mutagenesis screen and discovered that the ucp1 (you cannot pass) locus is responsible for the virulence of DC283. Analysis of the ucp1 gene sequence revealed similarities to the Rhs protein family. ucp1 gene is smaller than the genes encoding RHS/YD proteins and does not have a ligand binding YD repeat, but it has conserved 5′-cores, non-homologous 3′ ends, and it is a membrane bound protein. These structural similarities suggest enteric plant colonizers have the genetic ability to colonize insect hosts. Furthermore, the similarities between the ucp1 and rhs genes suggest that rhs-like genes have potential insecticidal activity (Stavrinides et al., 2010).